Microtubules are polarised α/β-tubulin-based polymers essential for cell viability, providing pushing and pulling forces, structural support or tracks for the transport of intracellular cargo (Goodson and Jonasson, 2018). α-tubulin is located at the so-called minus end and β-tubulin is exposed at the so-called plus end, which is typically more dynamic. This inherent microtubule polarity is important for cell polarity, as different motor proteins (Kinesins and Dynein) move cargo along microtubules in a specific direction – Dynein towards the minus end and most Kinesins towards the plus end. In neurons, most plus ends point away from the soma in axons (plus-end-out microtubules), while the microtubules in dendrites are either of mixed polarity or are predominantly minus-end-out (Hill et al., 2012; Kapitein and Hoogenraad, 2015; Kelliher et al., 2019; Tas et al., 2017). This difference between axons and dendrites is important for the correct distribution of cargo throughout the neuron (Harterink et al., 2018; Kapitein and Hoogenraad, 2015; Tas et al., 2017).

Within cells de novo assembly of new microtubules, that is microtubule nucleation, is kinetically unfavourable and is templated and catalysed by multi-protein γ-tubulin ring complexes (γ-TuRCs) (Tovey and Conduit, 2018). Knockdown of γ-TuRCs within model systems affects dynamic microtubules in all neuronal compartments (Nguyen et al., 2014; Ori-McKenney et al., 2012; Sánchez-Huertas et al., 2016; Yamada and Hayashi, 2019; Yau et al., 2014) and mutations in γ-TuRC genes have been linked to human neurodevelopmental disorders (Bahi-Buisson et al., 2014; Mitani et al., 2019; Poirier et al., 2013). γ-TuRCs are typically inactive until they are recruited to specific sites within cells, such as microtubule organising centres (MTOCs), the cytosol around mitotic chromatin, or the sides of pre-existing microtubules via binding to Augmin/HAUS complexes (Farache et al., 2018; Lin et al., 2015; Meunier and Vernos, 2016; Sanchez and Feldman, 2017; Teixido-Travesa et al., 2012). A range of MTOCs exist, including centrosomes, the Golgi apparatus and the nuclear envelope, and different cells use different MTOCs to help generate and organise their specific microtubule arrays (Sanchez and Feldman, 2017). γ-TuRC recruitment occurs via γ-TuRC ‘tethering proteins’, such as Drosophila Centrosomin (Cnn), that simultaneously bind to the γ-TuRC and a particular MTOC, and can also help activate the γ-TuRC (Tovey and Conduit, 2018).

Although γ-tubulin is important within neurons (Nguyen et al., 2014; Ori-McKenney et al., 2012; Sánchez-Huertas et al., 2016; Yamada and Hayashi, 2019; Yau et al., 2014), it remains unclear how microtubule nucleation is regulated. During early development of mammalian neurons, the centrosome within the soma nucleates microtubules (Stiess et al., 2010) that are severed and transported into neurites via motor-based microtubule sliding (Baas et al., 2005). Microtubule sliding is also important for axon outgrowth in Drosophila cultured neurons (Del Castillo et al., 2015; Lu et al., 2013), and for establishing microtubule polarity (del Castillo et al., 2015; Klinman et al., 2017; Rao et al., 2017; Yan et al., 2013; Zheng et al., 2008). Centrosomes are inactivated, however, at later developmental stages (Stiess et al., 2010) and are dispensable for neuronal development in both mammalian and fly neurons (Nguyen et al., 2011; Stiess et al., 2010). No other active MTOCs within the neuronal soma have been described. Nevertheless, microtubules continue to grow within the soma (Nguyen et al., 2011; Sánchez-Huertas et al., 2016), and in mammalian neurons this depends in part on the HAUS complex (Sánchez-Huertas et al., 2016), which is also important for microtubule growth within axons and dendrites (Cunha-Ferreira et al., 2018; Sánchez-Huertas et al., 2016). Some MTOCs have been identified within dendrites: the basal body, or its surrounding region, within the distal part of C. elegans ciliated neurons acts as an MTOC, as does a similar region within the URX non-ciliated neuron (Harterink et al., 2018); an MTOC made from endosomes that tracks the dendritic growth cone in C. elegans PVD neurons has recently been identified (Liang et al., 2020); and fragments of Golgi called Golgi outposts within the dendrites of Drosophila dendritic arborisation (da) neurons are thought to recruit γ-TuRCs and act as MTOCs (Ori-McKenney et al., 2012; Yalgin et al., 2015; Zhou et al., 2014).

Drosophila larval da neurons are a popular in vivo neuronal model (Jan and Jan, 2010). Ease of imaging and genetic manipulation coupled with the ability to examine different neuronal classes, each with a stereotypical dendritic branching pattern, make them a model of choice when examining microtubule organisation within neurons. Four classes exist, including class I neurons that are proprioceptive and have the simplest ‘comb-like’ dendritic branching pattern, and class IV neurons that are nociceptive and have the most elaborate branching pattern, tiling the surface of the larva (Figure 1—figure supplement 1; Grueber et al., 2002). In both neuronal classes, microtubules in axons are predominantly plus-end-out throughout development, but microtubule polarity in dendrites progressively becomes more minus-end-out as the neurons develop; at two days post larval hatching the majority of dynamic microtubules are minus-end-out (Hill et al., 2012; Stone et al., 2008). Microtubule polarity within da neurons can be disrupted when microtubule regulators are depleted (Mattie et al., 2010; Nguyen et al., 2014; Ori-McKenney et al., 2012; Rolls and Jegla, 2015; Sears and Broihier, 2016; Weiner et al., 2016; Yalgin et al., 2015; Zhou et al., 2014), but we lack a full understanding of how microtubule polarity is established and maintained.

Golgi outposts are thought to provide localised sites of microtubule nucleation within the dendrites of da neurons to help regulate dendritic outgrowth and microtubule polarity (Ori-McKenney et al., 2012; Yalgin et al., 2015; Zhou et al., 2014). Unlike the somatic Golgi, which comprises cis, medial, and trans compartments organised into stacks and ribbons (Kondylis and Rabouille, 2009; Nakamura et al., 1995), Golgi outposts can be either single- or multi-compartment units with multi-compartment units having a higher propensity to initiate microtubule growth (Zhou et al., 2014). Within class I da neurons, Golgi outpost-mediated nucleation is thought to be dependent on the γ-TuRC-tethering protein Cnn and to help restrict dendritic branching (Yalgin et al., 2015), while within class IV neurons Golgi outpost-mediated nucleation is believed to be dependent on Pericentrin-like protein (Plp) and to help promote dendritic branching (Ori-McKenney et al., 2012). There remains some uncertainty, however, about the role of Golgi outposts in microtubule nucleation, as a separate study that examined the localisation of ectopically expressed γ-tubulin-GFP concluded that γ-tubulin localises predominantly to dendritic branch points in a Golgi outpost-independent manner (Nguyen et al., 2014).

In this study, we initially aimed to identify sites of microtubule nucleation within Drosophila da neurons. We used endogenously tagged γ-tubulin-GFP as a proxy for γ-TuRCs and performed a detailed analysis of γ-tubulin localisation within both class I and class IV da neurons. We find variation between neuronal classes in how γ-tubulin localises within dendrites, but find that the most prominent localisation of γ-tubulin is at the cis-compartment of somatic Golgi stacks within all sensory neurons of the dorsal cluster. This asymmetric γ-tubulin-Golgi association is not dependent on either Cnn or Plp, suggesting another molecule must regulate γ-TuRC recruitment to the Golgi. Tracking of EB1-GFP comets within the soma suggests that microtubules are nucleated asymmetrically from the somatic Golgi. We find that these Golgi-derived microtubules initially grow preferentially towards the axon. During growth, they are also guided towards the axon and away from dendrites in a Kinesin-2-dependent manner, and they readily enter the axon contributing to plus-end-out microtubule polarity. In contrast, the relatively small number of microtubules that do grow towards a dendrite are normally excluded and this also depends upon Kinesin-2. After Kinesin-2 depletion, growing microtubules more readily approach and enter the dendrites causing a dramatic increase in the proportion of anterograde microtubules in proximal dendrites. This results in a reversal of overall microtubule polarity in proximal dendrites from predominantly minus-end-out to predominantly plus-end-out. We therefore propose that plus-end-associated Kinesin-2 guides growing microtubules towards the axon and away from dendrites along a polarised microtubule network within the soma, while at dendrite entry points Kinesin-2 generates a stalling force on growing microtubules when engaging dendritic microtubules of opposite polarity.

Understanding of how neuronal microtubules are generated and organised is a matter of much interest and importance. In this paper, we have used Drosophila da neurons as an in vivo model system to address these questions. Our results have led us to propose a model that describes how microtubules are generated and organised within the soma, which also impacts the microtubule populations within proximal axons and dendrites. This model comprises multiple elements. Firstly, the model states that microtubules are nucleated asymmetrically from the somatic Golgi, growing initially with a preference towards the axon and away from dendrites. Secondly, plus-end-associated Kinesin-2 helps guide these growing microtubules towards the axon and away from dendrites along a polarised network of microtubules within the soma. Thirdly, plus-end-associated Kinesin-2 also helps prevent any microtubules that do approach the entrance to a dendrite from entering. We propose that a combination of these mechanisms is essential to maintain minus-end-out polarity in proximal dendrites, a key feature that distinguishes dendrites from axons.

We have shown that endogenously tagged γ-tubulin23C-GFP, which is a proxy for the major catalysts of microtubule nucleation, γ-TuRCs, localises predominantly to the Golgi stacks within the soma of da neurons, and to a few Golgi outposts within the proximal branches of class IV da neurons. Our data show that the ability of γ-tubulin to localise to these Golgi structures does not depend upon either Cnn or Plp, both of which have been implicated in recruiting γ-TuRCs to Golgi outposts within dendrites of class I and class IV da neurons, respectively (Ori-McKenney et al., 2012; Yalgin et al., 2015). Thus, γ-tubulin, presumably in the form of γ-TuRCs, must be recruited to the Golgi by another tethering protein, perhaps by one of the large coiled-coil ‘Golgin’ proteins that project from specific parts of the Golgi (Munro, 2011).

After showing that γ-tubulin-GFP localised to the somatic Golgi, we tracked EB1-GFP comets from the Golgi and found that they grew with an initial directional preference towards the axon, suggestive of asymmetric nucleation. This analysis was more robust under normal conditions than after cooling-warming cycles, because more comets could be analysed (it was difficult to keep the imaging plane consistent during temperature shifting, due to temperature-induced glass expansion/contraction). Thus, while an equal number of Golgi-derived comets (4) grew towards and away from the axon after warming in Figure 6—Video 2, we believe this perceived lack of asymmetry is most likely due to chance, especially as a fraction of comets can grow in more random directions even under normal conditions (Figure 6C). We therefore propose that there is a genuine preference for growth of Golgi-derived microtubules towards the axon. This may be explained, at least in part, by the asymmetric localisation of γ-tubulin at the somatic Golgi stacks. Using markers of cis- and trans-Golgi, we have found that γ-tubulin localises preferentially to the cis-Golgi. This is also true in mammalian cells, where the somatic Golgi acts as an asymmetric MTOC in various cell types, such as migrating fibroblasts and different types of cultured mammalian cells in interphase (Rios, 2014). It has been proposed that asymmetric microtubule nucleation is due to the presence of CLASP proteins at the trans-Golgi, which have been suggested to bind and stabilise the plus ends of microtubules that are nucleated by γ-TuRCs at the cis-Golgi (Vinogradova et al., 2009). Whether this occurs in Drosophila neurons remains to be explored. Drosophila CLASP is known to be a plus-tip protein within axons (Beaven et al., 2015) and is required for axon guidance (Lee et al., 2004), but to our knowledge a role for CLASP at the Golgi in any Drosophila cell type has not yet been reported. In mammalian cells, directional microtubule nucleation is thought to be important for cell motility in fibroblasts (Efimov et al., 2007) and for polarised pseudopod formation during invasion of cancer cells (Wu et al., 2016). Our data suggest that asymmetric nucleation at the somatic Golgi in Drosophila da neurons may establish a polarised microtubule network within the soma and is important for ensuring that plus-end-out microtubules enter axons rather than dendrites.

It is possible that a fraction of comets that originate from the somatic Golgi represent microtubules that were re-growing after partial depolymerisation, where the re-growth event happened to overlap a Golgi stack. However, several pieces of evidence suggest that the majority of the Golgi-derived comets represent nucleation events. First, γ-tubulin accumulates strongly at the somatic Golgi and a high accumulation of γ-TuRCs at a particular site is normally indicative of a nucleation site, such as when γ-TuRCs accumulate at mitotic centrosomes. Second, we have found that comets emerge repeatedly from the Golgi in a similar direction, strongly suggestive of repeated nucleation events. Third, we believe it is unlikely that all of the comets emerging from the Golgi after a cooling-warming cycle represent microtubules that had not fully depolymerised and that just happened to re-grow close to the Golgi. Moreover, comets that emerged from the Golgi a significant amount of time after warming did not emerge in the same direction as those that had grown immediately after warming. This strongly suggests they are not the same microtubule re-growing after depolymerisation but represent new nucleation events. Direct imaging of microtubules could help assess the degree of overall microtubule depolymerisation induced by cooling, although cold-resistant microtubules that may remain after cooling could impede the ability to visualise depolymerisation of dynamic microtubules, especially if the dynamic microtubules had originally grown alongside the stable microtubules. Increasing the cooling time before inducing regrowth may cause the depolymerisation of all microtubules, and this could be tested by using fluorescent markers of microtubules or by fixing and immunostaining the samples. These experiments have limitations, however, as keeping the animal alive for long enough to depolymerise all microtubule may be challenging and fixing and staining experiments do not allow one to observe the same neuron before, during, and after cooling. In summary, when we consider all of our data as a whole, we believe the most parsimonious conclusion is that microtubules are nucleated asymmetrically from the somatic Golgi.

It has previously been shown that centrosomes in Drosophila neurons do not act as MTOCs (Nguyen et al., 2011), and our observations that EB1-GFP comets do not emerge radially from a single source support this conclusion. In other cell types the somatic Golgi can ‘compete’ with centrosomes for the ability to organise microtubules, where decreasing nucleation from one organelle increases nucleation at the other (Gavilan et al., 2018; Ríos et al., 2004; Wu et al., 2016). It is therefore possible that the lack of microtubule nucleation at centrosomes in Drosophila neurons enables microtubules to be nucleated at the Golgi. In mammalian neurons the centrosome is deactivated after the first few days in culture (Stiess et al., 2010). An intriguing possibility is that this deactivation allows the Golgi to organise microtubules, which may be necessary to ensure correct microtubule polarity during later neuronal development, although nucleation from the Golgi in mammalian neurons has yet to be reported.

Our analysis was carried out in third instar larvae where the neurons are fully developed and functional. Axons have already extended and made connections in the central nervous system and dendritic arbors are largely established. It is thus somewhat surprising that microtubules continue to be nucleated within the soma. It is possible that somatic microtubules are necessary for transport of cargos to axonal and dendritic entry sites. Given that Golgi-derived microtubules grow preferentially towards the axon and that somatic Golgi stacks are distributed throughout the soma, there is likely an overall microtubule polarity within the soma itself, with more plus ends pointing towards the axon. Even small biases in the orientation of microtubules within a network can be important for the polarised distribution of components, such as in Drosophila oocytes (Zimyanin et al., 2008). A similar bias within neuronal soma could be important for directing molecules into axons and dendrites. Perhaps this asymmetric microtubule network within the soma, as well as the microtubules within the axon, need constant renewal. There is evidence that molecular motors can damage microtubules (Triclin et al., 2018) and so the large amount of motor-driven transport that occurs within neurons may necessitate microtubule renewal from the soma.

A key feature of our model is the action of plus-end-associated Kinesin-2. Our Kap3 RNAi data suggest that Kinesin-2 is required to guide growing microtubules along a polarised microtubule network within the soma towards the axon and away from dendrites, although future experiments with dual-colour imaging of microtubules and EB1-GFP comets will help support this conclusion. Our data also suggest that plus-end-associated Kinesin-2 prevents outward growing plus ends of somatic microtubules from entering dendrites. Within the soma, we envisage that when a somatic microtubule approaches another somatic microtubule at an acute angle (<90°), it can readily turn and be guided along that microtubule (Figure 8A,C). Kinesin-2 mediated "collision resolution" events occur at dendritic branch points in class I da neurons to ensure growing microtubules turn towards the soma (Weiner et al., 2016) and turning events have been recapitulated in vitro (Chen et al., 2014; Doodhi et al., 2014). One study showed that the in vitro turning events also occured at obtuse angles (>90°), where the growing microtubule buckled to allow the plus end to remain in contact with the ‘rail’ microtubule. This could presumably also occur within the soma of da neurons. The frequency of guidance events, however, decreases significantly as the angle increases (Doodhi et al., 2014). In contrast to the microtubule collision events that will occur with variable angles within the neuronal soma, when outward growing somatic microtubules try to grow into a dendrite they will always encounter dendritic microtubules at angles close to 180° (Figure 8B,C), because most dendritic microtubules are oriented minus-end-out and also because of the limited space available at dendrite entry points. Kinesin-2 motors on the plus end of the outward growing somatic microtubules could engage with the shafts of the minus-end-out dendritic microtubules and generate a backwards force when attempting to ‘walk’ towards the plus end of the dendritic microtubule i.e. in a direction opposite to which the plus end of the somatic microtubule is attempting to grow. This backward force would presumably lead to stalling of the plus end and thus depolymerisation of the microtubule. It is also possible that Kinesin-2 motors slide the plus end along the dendritic microtubule back into the soma, but this sliding action requires the growing microtubule to buckle and thus requires sufficient space (Doodhi et al., 2014). Thus, buckling may be impeded at the narrow dendrite entry site anddepolymerisation favoured. Kinesin-2 motors should be sufficient to induce stalling of microtubule growth, because growing microtubules generate a force of ~3–4 pN (Dogterom and Yurke, 1997), while a single Kinesin-2 motor generates a force of ~5 pN (Schroeder et al., 2012). Depolymerisation upon stalling could be equivalent to how depolymerisation is induced when microtubules grow against immovable objects (Janson et al., 2003). Importantly, this model elegantly explains why microtubules from the soma are prevented from growing into the dendrites while dendritic microtubules can readily grow into the soma. Other models, such as if a high concentration of a microtubule depolymerase was present at dendritic entry sites, could not distinguish between these two events. In contrast, our model predicts that dendritic microtubules can grow readily into the soma because they do not frequently encounter a microtubule with opposite polarity, except for the occasional somatic microtubule attempting to grow into a dendrite.

Our model explains how minus-end-out microtubule polarity is maintained within proximal dendrites, but what is the origin of minus-end-out microtubules? These microtubules could have been pushed out from the soma minus end first by motor-based sliding, a mechanism known to promote initial axon outgrowth in cultured Drosophila neurons (Lu et al., 2013; Lu et al., 2015). Alternatively, or in conjunction with this, they could have been nucleated at sites within dendrites and then have grown back towards the soma. Retrograde microtubule growth events do occur within the dendrites of da neurons, and some of these growth events originate from Golgi outposts (Ori-McKenney et al., 2012; Yalgin et al., 2015; Zhou et al., 2014) and from early endosomes at branchpoints (Weiner et al., 2020). It was shown that γ-tubulin co-localises with early endosome markers at class I branchpoints and that this localisation relies on Wnt signalling proteins (Weiner et al., 2020). Our observation that endogenously-tagged γ-tubulin localises to branchpoints supports this conclusion. Moreover, it was recently shown that an endosome-based MTOC tracks the growing dendritic growth cone within C. elegans PVD neurons and nucleates microtubules that travel back towards the soma (Liang et al., 2020). The authors also suggest that a similar process may occur in newly developing Drosophila class I neurons. We also observe γ-tubulin within dendritic bubbles and dendritic stretches, but whether γ-tubulin is also recruited to these regions by endosomes, or by cytosolic accumulations of γ-TuRC tethering proteins such as Cnn or Plp, will need to be addressed in future studies. While our data suggest that γ-TuRCs are absent from the majority of Golgi outposts, they appear to be present at a few proximal Golgi outposts within class IV neurons. Moreover, microtubule growth events from Golgi outposts could be independent of γ-TuRCs; other non-γ-TuRC microtubule binding proteins can promote microtubule nucleation in vitro (Roostalu and Surrey, 2017), and ectopic Golgi outposts within the axons of nudE mutants promote microtubule growth but not via γ-TuRCs (Arthur et al., 2015; Yang and Wildonger, 2020). Thus, there are several potential mechanisms to generate minus-end-out microtubules within dendrites, that will need to be explored further in future.

It will be interesting in future to examine whether the plus ends of somatic microtubules can grow into dendrites more readily during early neuronal development, when there may be fewer minus-end-out microtubules within dendrites. If true, there must come a developmental point at which the balance shifts and microtubules are prevented from growing into dendrites. This shift could occur in several different ways, including upregulating the number of retrograde microtubules nucleated within dendrites, upregulating Kinesin-2, or changing the direction of microtubule nucleation within the soma. These are all interesting possibilities that are now open for investigation in the future.

All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figure 6 and 7.

1. 1. Arthur AL
   2. Yang SZ
   3. Abellaneda AM
   4. Wildonger J

   (2015) Dendrite arborization requires the dynein cofactor NudE

   Journal of Cell Science 128:2191–2201.

   https://doi.org/10.1242/jcs.170316

   • PubMed
   • Google Scholar
2. 1. Baas PW
   2. Karabay A
   3. Qiang L

   (2005) Microtubules cut and run

   Trends in Cell Biology 15:518–524.

   https://doi.org/10.1016/j.tcb.2005.08.004

   • PubMed
   • Google Scholar
3. 1. Bahi-Buisson N
   2. Poirier K
   3. Fourniol F
   4. Saillour Y
   5. Valence S
   6. Lebrun N
   7. Hully M
   8. Bianco CF
   9. Boddaert N
   10. Elie C
   11. Lascelles K
   12. Souville I
   13. Beldjord C
   14. Chelly J
   15. LIS-Tubulinopathies Consortium

   (2014) The wide spectrum of tubulinopathies: what are the key features for the diagnosis?

   Brain 137:1676–1700.

   https://doi.org/10.1093/brain/awu082

   • PubMed
   • Google Scholar
4. 1. Beaven R
   2. Dzhindzhev NS
   3. Qu Y
   4. Hahn I
   5. Dajas-Bailador F
   6. Ohkura H
   7. Prokop A

   (2015) Drosophila CLIP-190 and mammalian CLIP-170 display reduced microtubule plus end association in the nervous system

   Molecular Biology of the Cell 26:1491–1508.

   https://doi.org/10.1091/mbc.E14-06-1083

   • Google Scholar
5. 1. Bosc C
   2. Andrieux A
   3. Job D

   (2003) STOP proteins

   Biochemistry-Us 42:12125–12132.

   https://doi.org/10.1021/bi0352163

   • Google Scholar
6. 1. Broadie KS
   2. Bate M

   (1993) Development of the embryonic neuromuscular synapse of Drosophila Melanogaster

   The Journal of Neuroscience 13:144–166.

   https://doi.org/10.1523/JNEUROSCI.13-01-00144.1993

   • PubMed
   • Google Scholar
7. 1. Bucciarelli E
   2. Pellacani C
   3. Naim V
   4. Palena A
   5. Gatti M
   6. Somma MP

   (2009) Drosophila Dgt6 interacts with Ndc80, msps/XMAP215, and gamma-tubulin to promote kinetochore-driven MT formation

   Current Biology 19:1839–1845.

   https://doi.org/10.1016/j.cub.2009.09.043

   • PubMed
   • Google Scholar
8. 1. Chen Y
   2. Rolls MM
   3. Hancock WO

   (2014) An EB1-kinesin complex is sufficient to steer microtubule growth in vitro

   Current Biology 24:316–321.

   https://doi.org/10.1016/j.cub.2013.11.024

   • PubMed
   • Google Scholar
9. 1. Chen JV
   2. Buchwalter RA
   3. Kao LR
   4. Megraw TL

   (2017) A splice variant of centrosomin converts mitochondria to Microtubule-Organizing centers

   Current Biology 27:1928–1940.

   https://doi.org/10.1016/j.cub.2017.05.090

   • PubMed
   • Google Scholar
10. 1. Cunha-Ferreira I
    2. Chazeau A
    3. Buijs RR
    4. Stucchi R
    5. Will L
    6. Pan X
    7. Adolfs Y
    8. van der Meer C
    9. Wolthuis JC
    10. Kahn OI
    11. Schätzle P
    12. Altelaar M
    13. Pasterkamp RJ
    14. Kapitein LC
    15. Hoogenraad CC

    (2018) The HAUS complex is a key regulator of Non-centrosomal microtubule organization during neuronal development

    Cell Reports 24:791–800.

    https://doi.org/10.1016/j.celrep.2018.06.093

    • PubMed
    • Google Scholar
11. 1. Del Castillo U
    2. Lu W
    3. Winding M
    4. Lakonishok M
    5. Gelfand VI

    (2015) Pavarotti/MKLP1 regulates microtubule sliding and neurite outgrowth in Drosophila neurons

    Current Biology 25:200–205.

    https://doi.org/10.1016/j.cub.2014.11.008

    • PubMed
    • Google Scholar
12. 1. del Castillo U
    2. Winding M
    3. Lu W
    4. Gelfand VI

    (2015) Interplay between kinesin-1 and cortical dynein during axonal outgrowth and microtubule organization in Drosophila neurons

    eLife 4:e10140.

    https://doi.org/10.7554/eLife.10140

    • PubMed
    • Google Scholar
13. 1. Delphin C
    2. Bouvier D
    3. Seggio M
    4. Couriol E
    5. Saoudi Y
    6. Denarier E
    7. Bosc C
    8. Valiron O
    9. Bisbal M
    10. Arnal I
    11. Andrieux A

    (2012) MAP6-F is a temperature sensor that directly binds to and protects microtubules from cold-induced depolymerization

    Journal of Biological Chemistry 287:35127–35138.

    https://doi.org/10.1074/jbc.M112.398339

    • PubMed
    • Google Scholar
14. 1. Dogterom M
    2. Yurke B

    (1997) Measurement of the force-velocity relation for growing microtubules

    Science 278:856–860.

    https://doi.org/10.1126/science.278.5339.856

    • PubMed
    • Google Scholar
15. 1. Doodhi H
    2. Katrukha EA
    3. Kapitein LC
    4. Akhmanova A

    (2014) Mechanical and geometrical constraints control kinesin-based microtubule guidance

    Current Biology 24:322–328.

    https://doi.org/10.1016/j.cub.2014.01.005

    • PubMed
    • Google Scholar
16. 1. Efimov A
    2. Kharitonov A
    3. Efimova N
    4. Loncarek J
    5. Miller PM
    6. Andreyeva N
    7. Gleeson P
    8. Galjart N
    9. Maia AR
    10. McLeod IX
    11. Yates JR
    12. Maiato H
    13. Khodjakov A
    14. Akhmanova A
    15. Kaverina I

    (2007) Asymmetric CLASP-dependent nucleation of noncentrosomal microtubules at the trans-Golgi network

    Developmental Cell 12:917–930.

    https://doi.org/10.1016/j.devcel.2007.04.002

    • PubMed
    • Google Scholar
17. 1. Eisman RC
    2. Phelps MA
    3. Kaufman TC

    (2009) Centrosomin: a complex mix of long and short isoforms is required for centrosome function during early development in Drosophila melanogaster

    Genetics 182:979–997.

    https://doi.org/10.1534/genetics.109.103887

    • PubMed
    • Google Scholar
18. 1. Farache D
    2. Emorine L
    3. Haren L
    4. Merdes A

    (2018) Assembly and regulation of γ-tubulin complexes

    Open Biology 8:170266.

    https://doi.org/10.1098/rsob.170266

    • PubMed
    • Google Scholar
19. 1. Gavilan MP
    2. Gandolfo P
    3. Balestra FR
    4. Arias F
    5. Bornens M
    6. Rios RM

    (2018) The dual role of the centrosome in organizing the microtubule network in interphase

    EMBO reports 19:e45942.

    https://doi.org/10.15252/embr.201845942

    • Google Scholar
20. 1. Goodson HV
    2. Jonasson EM

    (2018) Microtubules and Microtubule-Associated proteins

    Cold Spring Harbor Perspectives in Biology 10:a022608.

    https://doi.org/10.1101/cshperspect.a022608

    • PubMed
    • Google Scholar
21. 1. Grueber WB
    2. Jan LY
    3. Jan Y-N

    (2002)

    Tiling of the Drosophila epidermis by multidendritic sensory neurons

    Development 129:2878.

    • Google Scholar
22. 1. Han C
    2. Jan LY
    3. Jan YN

    (2011) Enhancer-driven membrane markers for analysis of nonautonomous mechanisms reveal neuron-glia interactions in Drosophila

    PNAS 108:9673–9678.

    https://doi.org/10.1073/pnas.1106386108

    • PubMed
    • Google Scholar
23. 1. Harterink M
    2. Edwards SL
    3. de Haan B
    4. Yau KW
    5. van den Heuvel S
    6. Kapitein LC
    7. Miller KG
    8. Hoogenraad CC

    (2018) Local microtubule organization promotes cargo transport in C. elegans dendrites

    Journal of Cell Science 131:jcs.223107.

    https://doi.org/10.1242/jcs.223107

    • Google Scholar
24. 1. Hayward D
    2. Metz J
    3. Pellacani C
    4. Wakefield JG

    (2014) Synergy between multiple microtubule-generating pathways confers robustness to centrosome-driven mitotic spindle formation

    Developmental Cell 28:81–93.

    https://doi.org/10.1016/j.devcel.2013.12.001

    • PubMed
    • Google Scholar
25. 1. Hill SE
    2. Parmar M
    3. Gheres KW
    4. Guignet MA
    5. Huang Y
    6. Jackson FR
    7. Rolls MM

    (2012) Development of dendrite polarity in Drosophila neurons

    Neural Development 7:34.

    https://doi.org/10.1186/1749-8104-7-34

    • PubMed
    • Google Scholar
26. 1. Jan YN
    2. Jan LY

    (2010) Branching out: mechanisms of dendritic arborization

    Nature Reviews Neuroscience 11:316–328.

    https://doi.org/10.1038/nrn2836

    • PubMed
    • Google Scholar
27. 1. Janson ME
    2. de Dood ME
    3. Dogterom M

    (2003) Dynamic instability of microtubules is regulated by force

    Journal of Cell Biology 161:1029–1034.

    https://doi.org/10.1083/jcb.200301147

    • PubMed
    • Google Scholar
28. 1. Kapitein LC
    2. Hoogenraad CC

    (2015) Building the neuronal microtubule cytoskeleton

    Neuron 87:492–506.

    https://doi.org/10.1016/j.neuron.2015.05.046

    • PubMed
    • Google Scholar
29. 1. Kelliher MT
    2. Saunders HA
    3. Wildonger J

    (2019) Microtubule control of functional architecture in neurons

    Current Opinion in Neurobiology 57:39–45.

    https://doi.org/10.1016/j.conb.2019.01.003

    • PubMed
    • Google Scholar
30. 1. Klinman E
    2. Tokito M
    3. Holzbaur ELF

    (2017) CDK5-dependent activation of dynein in the axon initial segment regulates polarized cargo transport in neurons

    Traffic 18:808–824.

    https://doi.org/10.1111/tra.12529

    • PubMed
    • Google Scholar
31. 1. Kondylis V
    2. Rabouille C

    (2009) The golgi apparatus: lessons from Drosophila

    FEBS Letters 583:3827–3838.

    https://doi.org/10.1016/j.febslet.2009.09.048

    • PubMed
    • Google Scholar
32. 1. Lee H
    2. Engel U
    3. Rusch J
    4. Scherrer S
    5. Sheard K
    6. Van Vactor D

    (2004) The microtubule plus end tracking protein orbit/MAST/CLASP acts downstream of the tyrosine kinase abl in mediating axon guidance

    Neuron 42:913–926.

    https://doi.org/10.1016/j.neuron.2004.05.020

    • PubMed
    • Google Scholar
33. 1. Liang X
    2. Kokes M
    3. Fetter RD
    4. Sallee MD
    5. Moore AW
    6. Feldman JL
    7. Shen K

    (2020) Growth cone-localized microtubule organizing center establishes microtubule orientation in dendrites

    eLife 9:e56547.

    https://doi.org/10.7554/eLife.56547

    • PubMed
    • Google Scholar
34. 1. Lin TC
    2. Neuner A
    3. Schiebel E

    (2015) Targeting of γ-tubulin complexes to microtubule organizing centers: conservation and divergence

    Trends in Cell Biology 25:296–307.

    https://doi.org/10.1016/j.tcb.2014.12.002

    • PubMed
    • Google Scholar
35. 1. Lu W
    2. Fox P
    3. Lakonishok M
    4. Davidson MW
    5. Gelfand VI

    (2013) Initial neurite outgrowth in Drosophila neurons is driven by kinesin-powered microtubule sliding

    Current Biology 23:1018–1023.

    https://doi.org/10.1016/j.cub.2013.04.050

    • PubMed
    • Google Scholar
36. 1. Lu W
    2. Lakonishok M
    3. Gelfand VI

    (2015) Kinesin-1–powered microtubule sliding initiates axonal regeneration in Drosophila cultured neurons

    Molecular Biology of the Cell 26:1296–1307.

    https://doi.org/10.1091/mbc.E14-10-1423

    • Google Scholar
37. 1. Lucas EP
    2. Raff JW

    (2007) Maintaining the proper connection between the centrioles and the pericentriolar matrix requires Drosophila centrosomin

    Journal of Cell Biology 178:725–732.

    https://doi.org/10.1083/jcb.200704081

    • PubMed
    • Google Scholar
38. 1. Martinez-Campos M
    2. Basto R
    3. Baker J
    4. Kernan M
    5. Raff JW

    (2004) The Drosophila pericentrin-like protein is essential for cilia/flagella function, but appears to be dispensable for mitosis

    Journal of Cell Biology 165:673–683.

    https://doi.org/10.1083/jcb.200402130

    • PubMed
    • Google Scholar
39. 1. Mattie FJ
    2. Stackpole MM
    3. Stone MC
    4. Clippard JR
    5. Rudnick DA
    6. Qiu Y
    7. Tao J
    8. Allender DL
    9. Parmar M
    10. Rolls MM

    (2010) Directed microtubule growth, +TIPs, and kinesin-2 are required for uniform microtubule polarity in dendrites

    Current Biology 20:2169–2177.

    https://doi.org/10.1016/j.cub.2010.11.050

    • PubMed
    • Google Scholar
40. 1. Meunier S
    2. Vernos I

    (2016) Acentrosomal microtubule assembly in mitosis: the where, when, and how

    Trends in Cell Biology 26:80–87.

    https://doi.org/10.1016/j.tcb.2015.09.001

    • PubMed
    • Google Scholar
41. 1. Mitani T
    2. Punetha J
    3. Akalin I
    4. Pehlivan D
    5. Dawidziuk M
    6. Coban Akdemir Z
    7. Yilmaz S
    8. Aslan E
    9. Hunter JV
    10. Hijazi H
    11. Grochowski CM
    12. Jhangiani SN
    13. Karaca E
    14. Fatih JM
    15. Iwanowski P
    16. Gambin T
    17. Wlasienko P
    18. Goszczanska-Ciuchta A
    19. Bekiesinska-Figatowska M
    20. Hosseini M
    21. Arzhangi S
    22. Najmabadi H
    23. Rosenfeld JA
    24. Du H
    25. Marafi D
    26. Blaser S
    27. Teitelbaum R
    28. Silver R
    29. Posey JE
    30. Ropers H-H
    31. Gibbs RA
    32. Wiszniewski W
    33. Lupski JR
    34. Chitayat D
    35. Kahrizi K
    36. Gawlinski P

    (2019) Bi-allelic pathogenic variants in TUBGCP2 cause microcephaly and lissencephaly spectrum disorders

    The American Journal of Human Genetics 105:1005–1015.

    https://doi.org/10.1016/j.ajhg.2019.09.017

    • Google Scholar
42. 1. Munro S

    (2011) The golgin coiled-coil proteins of the golgi apparatus

    Cold Spring Harbor Perspectives in Biology 3:a005256.

    https://doi.org/10.1101/cshperspect.a005256

    • PubMed
    • Google Scholar
43. 1. Nakamura N
    2. Rabouille C
    3. Watson R
    4. Nilsson T
    5. Hui N
    6. Slusarewicz P
    7. Kreis TE
    8. Warren G

    (1995) Characterization of a cis-Golgi matrix protein, GM130

    The Journal of Cell Biology 131:1715–1726.

    https://doi.org/10.1083/jcb.131.6.1715

    • PubMed
    • Google Scholar
44. 1. Nguyen MM
    2. Stone MC
    3. Rolls MM

    (2011) Microtubules are organized independently from the centrosome in Drosophila neurons

    Neural Development 6:38.

    https://doi.org/10.1186/1749-8104-6-38

    • Google Scholar
45. 1. Nguyen MM
    2. McCracken CJ
    3. Milner ES
    4. Goetschius DJ
    5. Weiner AT
    6. Long MK
    7. Michael NL
    8. Munro S
    9. Rolls MM

    (2014) Γ-tubulin controls neuronal microtubule polarity independently of golgi outposts

    Molecular Biology of the Cell 25:2039–2050.

    https://doi.org/10.1091/mbc.e13-09-0515

    • PubMed
    • Google Scholar
46. 1. Ori-McKenney KM
    2. Jan LY
    3. Jan YN

    (2012) Golgi outposts shape dendrite morphology by functioning as sites of acentrosomal microtubule nucleation in neurons

    Neuron 76:921–930.

    https://doi.org/10.1016/j.neuron.2012.10.008

    • PubMed
    • Google Scholar
47. 1. Poirier K
    2. Lebrun N
    3. Broix L
    4. Tian G
    5. Saillour Y
    6. Boscheron C
    7. Parrini E
    8. Valence S
    9. Pierre BS
    10. Oger M
    11. Lacombe D
    12. Geneviève D
    13. Fontana E
    14. Darra F
    15. Cances C
    16. Barth M
    17. Bonneau D
    18. Bernadina BD
    19. N'Guyen S
    20. Gitiaux C
    21. Parent P
    22. des Portes V
    23. Pedespan JM
    24. Legrez V
    25. Castelnau-Ptakine L
    26. Nitschke P
    27. Hieu T
    28. Masson C
    29. Zelenika D
    30. Andrieux A
    31. Francis F
    32. Guerrini R
    33. Cowan NJ
    34. Bahi-Buisson N
    35. Chelly J

    (2013) Mutations in TUBG1, DYNC1H1, KIF5C and KIF2A cause malformations of cortical development and microcephaly

    Nature Genetics 45:639–647.

    https://doi.org/10.1038/ng.2613

    • Google Scholar
48. 1. Rao AN
    2. Patil A
    3. Black MM
    4. Craig EM
    5. Myers KA
    6. Yeung HT
    7. Baas PW

    (2017) Cytoplasmic dynein transports axonal microtubules in a Polarity-Sorting manner

    Cell Reports 19:2210–2219.

    https://doi.org/10.1016/j.celrep.2017.05.064

    • PubMed
    • Google Scholar
49. 1. Ríos RM
    2. Sanchís A
    3. Tassin AM
    4. Fedriani C
    5. Bornens M

    (2004) GMAP-210 recruits gamma-tubulin complexes to cis-Golgi membranes and is required for golgi ribbon formation

    Cell 118:323–335.

    https://doi.org/10.1016/j.cell.2004.07.012

    • PubMed
    • Google Scholar
50. 1. Rios RM

    (2014) The centrosome–Golgi apparatus nexus

    Philosophical Transactions of the Royal Society B: Biological Sciences 369:20130462.

    https://doi.org/10.1098/rstb.2013.0462

    • Google Scholar
51. 1. Rolls MM
    2. Jegla TJ

    (2015) Neuronal polarity: an evolutionary perspective

    Journal of Experimental Biology 218:572–580.

    https://doi.org/10.1242/jeb.112359

    • PubMed
    • Google Scholar
52. 1. Roostalu J
    2. Surrey T

    (2017) Microtubule nucleation: beyond the template

    Nature Reviews Molecular Cell Biology 18:702–710.

    https://doi.org/10.1038/nrm.2017.75

    • Google Scholar
53. 1. Roubin R
    2. Acquaviva C
    3. Chevrier V
    4. Sedjai F
    5. Zyss D
    6. Birnbaum D
    7. Rosnet O

    (2013) Myomegalin is necessary for the formation of centrosomal and Golgi-derived microtubules

    Biology Open 2:238–250.

    https://doi.org/10.1242/bio.20123392

    • Google Scholar
54. 1. Sanchez AD
    2. Feldman JL

    (2017) Microtubule-organizing centers: from the centrosome to non-centrosomal sites

    Current Opinion in Cell Biology 44:93–101.

    https://doi.org/10.1016/j.ceb.2016.09.003

    • PubMed
    • Google Scholar
55. 1. Sánchez-Huertas C
    2. Freixo F
    3. Viais R
    4. Lacasa C
    5. Soriano E
    6. Lüders J

    (2016) Non-centrosomal nucleation mediated by augmin organizes microtubules in post-mitotic neurons and controls axonal microtubule polarity

    Nature Communications 7:12187.

    https://doi.org/10.1038/ncomms12187

    • PubMed
    • Google Scholar
56. 1. Schroeder HW
    2. Hendricks AG
    3. Ikeda K
    4. Shuman H
    5. Rodionov V
    6. Ikebe M
    7. Goldman YE
    8. Holzbaur EL

    (2012) Force-dependent detachment of kinesin-2 biases track switching at cytoskeletal filament intersections

    Biophysical Journal 103:48–58.

    https://doi.org/10.1016/j.bpj.2012.05.037

    • PubMed
    • Google Scholar
57. 1. Sears JC
    2. Broihier HT

    (2016) FoxO regulates microtubule dynamics and polarity to promote dendrite branching in Drosophila sensory neurons

    Developmental Biology 418:40–54.

    https://doi.org/10.1016/j.ydbio.2016.08.018

    • Google Scholar
58. 1. Sepp KJ
    2. Auld VJ

    (2003) Reciprocal interactions between neurons and Glia are required for Drosophila peripheral nervous system development

    The Journal of Neuroscience 23:8221–8230.

    https://doi.org/10.1523/JNEUROSCI.23-23-08221.2003

    • PubMed
    • Google Scholar
59. 1. Stiess M
    2. Maghelli N
    3. Kapitein LC
    4. Gomis-Rüth S
    5. Wilsch-Bräuninger M
    6. Hoogenraad CC
    7. Tolić-Nørrelykke IM
    8. Bradke F

    (2010) Axon extension occurs independently of centrosomal microtubule nucleation

    Science 327:704–707.

    https://doi.org/10.1126/science.1182179

    • PubMed
    • Google Scholar
60. 1. Stone MC
    2. Roegiers F
    3. Rolls MM

    (2008) Microtubules have opposite orientation in axons and dendrites of Drosophila Neurons

    Molecular Biology of the Cell 19:4122–4129.

    https://doi.org/10.1091/mbc.e07-10-1079

    • Google Scholar
61. 1. Tas RP
    2. Chazeau A
    3. Cloin BMC
    4. Lambers MLA
    5. Hoogenraad CC
    6. Kapitein LC

    (2017) Differentiation between oppositely oriented microtubules controls polarized neuronal transport

    Neuron 96:1264–1271.

    https://doi.org/10.1016/j.neuron.2017.11.018

    • PubMed
    • Google Scholar
62. 1. Teixido-Travesa N
    2. Roig J
    3. Luders J

    (2012) The where, when and how of microtubule nucleation - one ring to rule them all

    Journal of Cell Science 125:4445–4456.

    https://doi.org/10.1242/jcs.106971

    • Google Scholar
63. 1. Torosantucci L
    2. De Luca M
    3. Guarguaglini G
    4. Lavia P
    5. Degrassi F

    (2008) Localized RanGTP accumulation promotes microtubule nucleation at Kinetochores in somatic mammalian cells

    Molecular Biology of the Cell 19:1873–1882.

    https://doi.org/10.1091/mbc.e07-10-1050

    • PubMed
    • Google Scholar
64. 1. Torres IL
    2. Rosa-Ferreira C
    3. Munro S

    (2014) The arf family G protein Arl1 is required for secretory granule biogenesis in Drosophila

    Journal of Cell Science 127:2151–2160.

    https://doi.org/10.1242/jcs.122028

    • PubMed
    • Google Scholar
65. 1. Tovey CA
    2. Tubman CE
    3. Hamrud E
    4. Zhu Z
    5. Dyas AE
    6. Butterfield AN
    7. Fyfe A
    8. Johnson E
    9. Conduit PT

    (2018) γ-TuRC heterogeneity revealed by analysis of Mozart1

    Current Biology 28:2314–2323.

    https://doi.org/10.1016/j.cub.2018.05.044

    • PubMed
    • Google Scholar
66. 1. Tovey CA
    2. Conduit PT

    (2018) Microtubule nucleation by γ-tubulin complexes and beyond

    Essays in Biochemistry 62:765–780.

    https://doi.org/10.1042/EBC20180028

    • PubMed
    • Google Scholar
67. Preprint

    1. Triclin S
    2. Inoue D
    3. Gaillard J
    4. Htet ZM
    5. Santis MD
    6. Portran D
    7. Derivery E
    8. Aumeier C
    9. Schaedel L
    10. John K

    (2018) Self-repair protects microtubules from their destruction by molecular motors

    bioRxiv.

    https://doi.org/10.1101/499020

    • Google Scholar
68. 1. Vinogradova T
    2. Miller PM
    3. Kaverina I

    (2009) Microtubule network asymmetry in motile cells: role of Golgi-derived array

    Cell Cycle 8:2168–2174.

    https://doi.org/10.4161/cc.8.14.9074

    • PubMed
    • Google Scholar
69. 1. Wang Z
    2. Wu T
    3. Shi L
    4. Zhang L
    5. Zheng W
    6. Qu JY
    7. Niu R
    8. Qi RZ

    (2010) Conserved motif of CDK5RAP2 mediates its localization to centrosomes and the golgi complex

    Journal of Biological Chemistry 285:22658–22665.

    https://doi.org/10.1074/jbc.M110.105965

    • PubMed
    • Google Scholar
70. 1. Weiner AT
    2. Lanz MC
    3. Goetschius DJ
    4. Hancock WO
    5. Rolls MM

    (2016) Kinesin-2 and apc function at dendrite branch points to resolve microtubule collisions

    Cytoskeleton 73:35–44.

    https://doi.org/10.1002/cm.21270

    • PubMed
    • Google Scholar
71. 1. Weiner AT
    2. Seebold DY
    3. Torres-Gutierrez P
    4. Folker C
    5. Swope RD
    6. Kothe GO
    7. Stoltz JG
    8. Zalenski MK
    9. Kozlowski C
    10. Barbera DJ
    11. Patel MA
    12. Thyagarajan P
    13. Shorey M
    14. Nye DMR
    15. Keegan M
    16. Behari K
    17. Song S
    18. Axelrod JD
    19. Rolls MM

    (2020) Endosomal wnt signaling proteins control microtubule nucleation in dendrites

    PLOS Biology 18:e3000647.

    https://doi.org/10.1371/journal.pbio.3000647

    • PubMed
    • Google Scholar
72. 1. Wu J
    2. de Heus C
    3. Liu Q
    4. Bouchet BP
    5. Noordstra I
    6. Jiang K
    7. Hua S
    8. Martin M
    9. Yang C
    10. Grigoriev I
    11. Katrukha EA
    12. Altelaar AFM
    13. Hoogenraad CC
    14. Qi RZ
    15. Klumperman J
    16. Akhmanova A

    (2016) Molecular pathway of microtubule organization at the golgi apparatus

    Developmental Cell 39:44–60.

    https://doi.org/10.1016/j.devcel.2016.08.009

    • Google Scholar
73. 1. Yadav S
    2. Younger SH
    3. Zhang L
    4. Thompson-Peer KL
    5. Li T
    6. Jan LY
    7. Jan YN

    (2019) Glial ensheathment of the somatodendritic compartment regulates sensory neuron structure and activity

    PNAS 116:5126–5134.

    https://doi.org/10.1073/pnas.1814456116

    • PubMed
    • Google Scholar
74. 1. Yalgin C
    2. Ebrahimi S
    3. Delandre C
    4. Yoong LF
    5. Akimoto S
    6. Tran H
    7. Amikura R
    8. Spokony R
    9. Torben-Nielsen B
    10. White KP
    11. Moore AW

    (2015) Centrosomin represses dendrite branching by orienting microtubule nucleation

    Nature Neuroscience 18:1437–1445.

    https://doi.org/10.1038/nn.4099

    • Google Scholar
75. 1. Yamada M
    2. Hayashi K

    (2019) Microtubule nucleation in the cytoplasm of developing cortical neurons and its regulation by brain-derived neurotrophic factor

    Cytoskeleton 76:339–345.

    https://doi.org/10.1002/cm.21550

    • PubMed
    • Google Scholar
76. 1. Yan J
    2. Chao DL
    3. Toba S
    4. Koyasako K
    5. Yasunaga T
    6. Hirotsune S
    7. Shen K

    (2013) Kinesin-1 regulates dendrite microtubule polarity in Caenorhabditis elegans

    eLife 2:e00133.

    https://doi.org/10.7554/eLife.00133

    • PubMed
    • Google Scholar
77. 1. Yang SZ
    2. Wildonger J

    (2020) Golgi outposts locally regulate microtubule orientation in neurons but are not required for the overall polarity of the dendritic cytoskeleton

    Genetics 215:435–447.

    https://doi.org/10.1534/genetics.119.302979

    • PubMed
    • Google Scholar
78. 1. Yau KW
    2. van Beuningen SFB
    3. Cunha-Ferreira I
    4. Cloin BMC
    5. van Battum EY
    6. Will L
    7. Schätzle P
    8. Tas RP
    9. van Krugten J
    10. Katrukha EA
    11. Jiang K
    12. Wulf PS
    13. Mikhaylova M
    14. Harterink M
    15. Pasterkamp RJ
    16. Akhmanova A
    17. Kapitein LC
    18. Hoogenraad CC

    (2014) Microtubule Minus-End binding protein CAMSAP2 controls axon specification and dendrite development

    Neuron 82:1058–1073.

    https://doi.org/10.1016/j.neuron.2014.04.019

    • Google Scholar
79. 1. Zheng Y
    2. Wildonger J
    3. Ye B
    4. Zhang Y
    5. Kita A
    6. Younger SH
    7. Zimmerman S
    8. Jan LY
    9. Jan YN

    (2008) Dynein is required for polarized dendritic transport and uniform microtubule orientation in axons

    Nature Cell Biology 10:1172–1180.

    https://doi.org/10.1038/ncb1777

    • Google Scholar
80. 1. Zhou W
    2. Chang J
    3. Wang X
    4. Savelieff MG
    5. Zhao Y
    6. Ke S
    7. Ye B

    (2014) GM130 is required for compartmental organization of dendritic golgi outposts

    Current Biology 24:1227–1233.

    https://doi.org/10.1016/j.cub.2014.04.008

    • PubMed
    • Google Scholar
81. 1. Zimyanin VL
    2. Belaya K
    3. Pecreaux J
    4. Gilchrist MJ
    5. Clark A
    6. Davis I
    7. St Johnston D

    (2008) In vivo imaging of oskar mRNA transport reveals the mechanism of posterior localization

    Cell 134:843–853.

    https://doi.org/10.1016/j.cell.2008.06.053

    • Google Scholar

Author details

1. Amrita Mukherjee

   Department of Zoology, University of Cambridge, Cambridge, United Kingdom

   Contribution

   Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing

   Competing interests

   No competing interests declared

2. Paul S Brooks

   Department of Zoology, University of Cambridge, Cambridge, United Kingdom

   Contribution

   Resources, Data curation, Formal analysis, Investigation, Methodology

   Competing interests

   No competing interests declared

3. Fred Bernard

   Université de Paris, CNRS, Institut Jacques Monod, Paris, France

   Contribution

   Resources

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-7919-253X

4. Antoine Guichet

   Université de Paris, CNRS, Institut Jacques Monod, Paris, France

   Contribution

   Resources, Data curation, Formal analysis, Investigation, Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-7216-1944

5. Paul T Conduit

   1. Department of Zoology, University of Cambridge, Cambridge, United Kingdom
   2. Université de Paris, CNRS, Institut Jacques Monod, Paris, France

   Contribution

   Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing

   For correspondence

   paul.conduit@ijm.fr

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-7822-1191

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